Funding and support were acquired through NIH grant K22 AI153678-01 and Rutgers School of Graduate Studies. We thank Fawad Yousufzai and Luke Fritzky from the Rutgers Biomedical Health Sciences Cellular Imaging and Histology Core for their work and expertise in obtaining immunofluorescent images.

All methods described in this protocol have been approved by the Institutional Animal Care and Use Committee (IACUC) of Rutgers Biomedical Health Sciences.
1. Label spores with intracellular fluorescence
<ol>
	<li>Stain 1 x 106 spores in carboxyfluorescein succinimidyl ester (CFSE), Cell Tracker Orange, or Cell Tracker Red (or intracellular dye of choice) at 5 &#181;M for 30 min at 30 &#176;C, then wash with PBS, and spin the spores at 12,000 x g. Repeat this wash once.</li>
	<li>Prepare the spore inoculum at the appropriate concentration so that 25 &#181;L contains the appropriate spore dose for one mouse.</li>
</ol>
2. Inoculating mouse with spores through aspiration inhalation using isoflurane anesthesia
CAUTION: Isoflurane is a volatile anesthetic agent and must be used within a ducted biosafety cabinet or fume hood.
<ol>
	<li>To prepare the isoflurane exposure jar, place a folded napkin into a 1 L screw-top Nalgene jar and place a plastic mesh circle over the napkin as a platform for the mice. Add 2 mL of isoflurane to the napkin, close the jar, and wait 1-2 min for the gas to equilibrate in the container.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67556/67556fig01.jpg" /> 
Figure 1: Apparatus for isoflurane open drop method of spore inoculation. A folded napkin is placed in the bottom of a 1 L screw top jar, and then a circular plastic mesh is inserted to act as a platform for the mice. <a href="https://www.jove.com/files/ftp_upload/67556/67556fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="2">
	<li>Prior to exposing the mouse to anesthesia, prepare a pipet with 25 &#181;L of the inoculum. Place the mouse in the isoflurane container and tilt the container to monitor for loss of the righting reflex, where the mouse has lost consciousness and is upside down. This usually takes about 10 s.</li>
	<li>Monitor the respiration rate (RR) until it slows to a consistent 50-80 breaths per minute (bpm), about 50% of resting RR. This usually takes another 10 s. At this time, remove the mouse from the isoflurane jar. 
	NOTE: This is a high concentration of isoflurane, which will produce rapid induction of anesthesia, and the mouse must be very closely monitored. Lethal isoflurane overdose is a risk, as the concentration of isoflurane can be variable, and the dose-response curve to isoflurane is very steep. If the breathing rate dips below 50 bpm and/or is irregular, remove the mouse from isoflurane, allow it to recover for 3-5 min, and try again (a maximum of two repeated attempts).</li>
	<li>Confirm&#160;the anesthetic depth by lack of response to toe pinch. Then, position the mouse in a supine position in one hand, and allow the mouse to take 3-5 regular gasping breaths with increasing rate (beginning at 50-60 bpm) and vigor. As this begins, place the pipet tip at the posterior of the mouse oropharynx and release the inoculum during an inhalation. 
	NOTE: These gasps through the mouth jolt the head inferiorly due to the recruitment of accessory muscles for breathing. When the pipet is inserted during these gasps, the mouth should open with each gasp. If the mouth does not open, the mouse is insufficiently anesthetized and will likely not inhale contents from the oropharynx. The pipet should depress the tongue, pushing it towards the bottom and front of the mouth to prevent swallowing of the inoculum. When the pipet is in the mouth and the mouse is adequately anesthetized, the mouse will open its mouth with each gasp.</li>
	<li>With the hand and thumb holding the mouse, feel for crackles on the posterior and anterior aspects of the mouse thorax to confirm inhalation of the inoculum.</li>
</ol>
3. Euthanasia of mice
<ol>
	<li>Inject the mouse intraperitoneally with a cocktail of 200 mg/kg ketamine and 24 mg/kg xylazine 10 min prior to the sacrifice time point.</li>
	<li>Wait 10 min post-injection until the mouse enters a surgical plane of anesthesia, confirmed by lack of response to toe pinch.</li>
</ol>
4. Perfusion of lungs with PBS and formalin
<ol>
	<li>Spray the anesthetized mouse with 70% ethanol. Use scissors to snip back the skin of the mouse and pull apart the skin to expose the peritoneum on all sides. Pull the skin from the mouse&#39;s superior half over its head, pulling out the arms from the skin.</li>
	<li>Cut the peritoneal membrane at the sternum and along the inferior aspect of the ribcage, exposing the peritoneum. Displace the liver inferiorly, revealing the diaphragm. Use scissors to carefully puncture the diaphragm away from the lung parenchyma to avoid puncturing a hole in the lungs themselves. 
	NOTE: Inadvertently puncturing the lungs will make subsequent air inflation impossible.</li>
	<li>After the diaphragm is punctured, air enters the thorax, collapsing the lungs. Cut the ribcage on the left side to reveal the heart. Inject 5-10 mL of PBS into the right ventricle with a 30 G needle at a rate of 1 mL per 5 s (to avoid damaging vessels), rapidly whitening the lungs. 
	CAUTION: Formalin is a hazardous volatile substance that should be handled within a ducted biosafety cabinet or fume hood.</li>
	<li>When finished, remove the needle and use a new needle to inject 5 mL of 10% neutral-buffered formalin (NBF) into the right ventricle at the same rate.</li>
</ol>
5. Air-inflating the perfused lung
<ol>
	<li>Remove the anterior half of the ribcage. To expose the trachea, cut the superior ribs and collarbones to the right and left of the neck. Take care to avoid cutting near the midline where the trachea will lie. With forceps, clasp the remaining top ribs and collarbones on the neck midline and pull superiorly until the trachea is exposed.</li>
	<li>Prepare a suture thread of 10 cm in length, pull it under the trachea, and pre-tie a loose knot at the inferior end of the exposed trachea. Prepare a 1 mL syringe of air with an 18 G catheter. Use an 18 G needle to make a hole in the trachea at the superior end of the exposed region.</li>
	<li>Place the catheter into that hole, ensuring a relatively tight fit to prevent air from escaping. Slowly inject the 1 mL of air into the lungs over the course of 10 s, watching for lung inflation of all lobes. Full inflation results in the lungs wrapping slightly around the heart and filling the volume they occupy in the unpunctured diaphragm.</li>
	<li>Pull the knot tight around the trachea, and remove the catheter. Hold the trachea and use blunt scissors to remove the lungs from the mouse, being careful not to puncture the lung.</li>
</ol>
6. Immersion fixation and dehydration
<ol>
	<li>Place the lungs on the top edge of a 50 mL conical tube filled with 20 mL of 10% NBF. Keep the suture threads outside of the conical, screw down the cap, and invert the conical so that the lungs are suspended upside down in 20 mL of NBF. Place it at 4 &#176;C for 24-48 h.</li>
	<li>Rinse the lungs in PBS, then place in a 30% sucrose-PBS (w/v) solution for 72-96 h at 4 &#176;C to dehydrate the lungs in preparation for cryopreservation. Remove the lungs from sucrose, place the lungs in a cryomold with optimal cutting temperature (OCT) medium, and freeze the specimen at -80 &#176;C.</li>
</ol>
7. Cryosectioning
<ol>
	<li>Equilibrate the specimens in OCT cryoblocks to the -20 &#176;C temperature of the cryostat for 1 h prior to sectioning. Section the blocks at thicknesses of 20-100 &#181;m. 
	NOTE: Air inflation can lead to increased fragility of the specimen, and cutting at increased thickness can prevent tissue breakage during sectioning.</li>
	<li>Collect sections onto glass slides and let dry for 30 min to 1 h to ensure tissue adherence to the slide. 
	NOTE: Slides may be held at -80 &#176;C at this stage.</li>
</ol>
8. Slide preparation and blocking
<ol>
	<li>Place slides in PBS bath (in a dish or Coplin jar) for 30 min at room temperature (RT) to remove OCT from the slides. 
	NOTE: Thicker sections (40-100 &#181;m) may not adhere to glass slides as well as thinner sections, so it is best to keep them flat and upright in a dish rather than placed on their side in a Coplin jar.</li>
	<li>Dry the slides after OCT has dissolved away from the tissue. Use a wipe to dry excess droplets from the slide&#39;s perimeter around the tissue specimen. </li>
	<li>Use a hydrophobic Pap pen to draw a perimeter around the specimen, and let it dry for 5 min.</li>
	<li>Prepare a blocking buffer of animal-free blocking solution with 0.3% Tween-20 and 1:100 Fc Block in 300 mL per slide. 
	NOTE: For intracellular targets, 1% Tween-20 may be used. A volume of 300 &#181;L is usually sufficient, but the necessary volume to cover the tissue fully depends on the perimeter size of the hydrophobic marker. The tissue should not dry from this point onwards.</li>
	<li>Leave the blocking solution on the slides for 1 h at RT.</li>
</ol>
9. Immunostaining
<ol>
	<li>Wash the slide by pipetting off the fluid and adding 300-500 &#181;L of PBS.&#160;Repeat the wash once.</li>
	<li>Prepare primary AlexaFluor-647 conjugated rat anti-mouse EpCAM (clone G8.8) antibody (airway epithelial marker) in animal-free blocker solution with 0.33% Tween-20 (or 1% for intracellular targets). Add 300 &#181;L per slide of this solution and stain overnight at 4 &#176;C. 
	NOTE: Antibodies should be tested empirically for staining concentration, time, and temperature. Thin sections (8-20 &#181;m) are stained for 30 min at 37 &#176;C and thicker sections (20-100 &#181;m) at 4 &#176;C overnight using 1:100 dilutions for most antibodies. If using unconjugated antibodies, wash 2x (as above) and apply a secondary fluorescent antibody for an empirically determined time and temperature. Generally, secondary antibody staining is done at RT for 1 h at 1:1000 dilution.</li>
	<li>Remove the antibody solution from the specimen, and wash the slide with 300 mL of PBS for 5 min. Repeat this wash once. Dry the slides, removing all excess fluid from the specimen and surrounding glass.</li>
	<li>Add one drop of RT soft set mounting media (e.g., SlowFade Glass) to each piece of tissue. Take care to avoid bubbles by letting the bottle settle while upside down and slowly dropping each drop onto the tissues. Place an appropriately sized cover slip to extend beyond the sample and hydrophobic marker perimeter.</li>
</ol>
10. Imaging via fluorescent microscopy
<ol>
	<li>Use a multichannel fluorescent microscope to scan whole lung sections in an unbiased manner using an objective with sufficient resolution to resolve individual spores and host cells (e.g., Zeiss Axioscan 7 using 20x oil objective with NA = 0.8 or similar). 
	NOTE: Ensure that laser power and PMT voltage gain are set to maximize the signal-to-noise ratios of the target over the background determined with a fluorescence minus one (FMO) control.</li>
	<li>Collect sufficient image tiles to capture the entire lung lobe (e.g., ~200 tiles of size 400 &#956;m x 400 &#956;m). Use the microscope software program (e.g., Zeiss ZEN 3.7 or similar) to export the merged tile image for downstream processing.</li>
</ol>
11. Spatial analysis via QuPath
<ol>
	<li>In QuPath, a free open-source software14, create a project file and add the fluorescent images to the file.</li>
	<li>Classify the EpCAM+ pixels as epithelium by clicking the top menu option Classify, then Pixel classification &#62; Create Thresholder. Select the resolution, channel, smoothing sigma, and threshold value that best identifies the target region. Save the classifier under a unique name, and select Create Objects.</li>
	<li>In the new Create objects window, select New object type as Annotation. For lung epithelium, set the minimum object size and minimum hole size to 100 mm2. There is no need to select Split Objects here. After selecting OK, the annotations will be created, and they can be modified by eye using the Brush tool in the upper left-hand area.</li>
	<li>To classify spores, repeat the steps from 11.3-11.4 using the channel in which the spores were captured and make the new object type Detection rather than Annotation. Set minimum object size and minimum hole size to 0 mm2, and select Split Objects.</li>
	<li>To conduct spatial analysis of the spores, select the top menu option Analyze &#62; Spatial Analysis &#62; Calculate signed distance to annotations 2D. The distances to EpCAM+ epithelium will be calculated for each detected spore. Export these via the top menu option Measure &#62; Export Measurements.</li>
</ol>

Preserving Fungal Spore Localization in Lungs for Fluorescent Microscopy Analysis

<ol>
	<li>American Society for Microbiology. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=One+Health:+Fungal+Pathogens+of+Humans,+Animals,+and+Plants.">One Health: Fungal Pathogens of Humans, Animals, and Plants.</a> Report on an American Academy of Microbiology Colloquium. , (2019).</li><li>Crossen, A. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+airway+epithelium+responses+to+invasive+fungal+infections:+A+critical+partner+in+innate+immunity.">Human airway epithelium responses to invasive fungal infections: A critical partner in innate immunity.</a> J Fungi (Basel). 9 (1), 40 (2022).</li><li>Yamamoto, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Particle-size+distributions+and+seasonal+diversity+of+allergenic+and+pathogenic+fungi+in+outdoor+air.">Particle-size distributions and seasonal diversity of allergenic and pathogenic fungi in outdoor air.</a> ISME J. 6 (10), 1801-1811 (2012).</li><li>Thakur, A. K., Kaundle, B., Singh, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Mucoadhesive Drug Delivery Systems in Respiratory Diseases.Targeting Chronic Inflammatory Lung Diseases Using Advanced Drug Delivery Systems. , (2020).</li><li>Darquenne, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aerosol+deposition+in+health+and+disease.">Aerosol deposition in health and disease.</a> J Aerosol Med Pulm Drug Deliv. 25 (3), 140-147 (2012).</li><li>Kradin, R. L., Mark, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pathology+of+pulmonary+disorders+due+to+Aspergillus+spp.">The pathology of pulmonary disorders due to Aspergillus spp.</a> Arch Pathol Lab Med. 132 (4), 606-614 (2008).</li><li>Karnak, D., Avery, R. K., Gildea, T. R., Sahoo, D., Mehta, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endobronchial+fungal+disease:+An+under-recognized+entity.">Endobronchial fungal disease: An under-recognized entity.</a> Respiration. 74 (1), 88-104 (2006).</li><li>Amich, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+light+sheet+fluorescence+microscopy+of+lungs+to+dissect+local+host+immune-Aspergillus+fumigatus+interactions.">Three-dimensional light sheet fluorescence microscopy of lungs to dissect local host immune-Aspergillus fumigatus interactions.</a> mBio. 11 (1), e02752-e02819 (2020).</li><li>Wiesner, D. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Club+cell+TRPV4+serves+as+a+damage+sensor+driving+lung+allergic+inflammation.">Club cell TRPV4 serves as a damage sensor driving lung allergic inflammation.</a> Cell Host Microbe. 27 (4), 614-628.e6 (2020).</li><li>Evans, S. E., Hahn, P. Y., McCann, F., Kottom, T. J., Pavlovic', Z. V., Limper, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pneumocystis+cell+wall+&#946;-glucans+stimulate+alveolar+epithelial+cell+chemokine+generation+through+nuclear+factor-&#954;B-dependent+mechanisms.">Pneumocystis cell wall &#946;-glucans stimulate alveolar epithelial cell chemokine generation through nuclear factor-&#954;B-dependent mechanisms.</a> Am J Respir Cell Mol Biol. 32 (6), 490-497 (2005).</li><li>Bauer, C., Krueger, M., Lamm, W. J. E., Glenny, R. W., Beichel, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=lapdMouse:+associating+lung+anatomy+with+local+particle+deposition+in+mice.">lapdMouse: associating lung anatomy with local particle deposition in mice.</a> J Appl Physiol. 128 (2), 309-323 (2020).</li><li>Srirama, P. K., Wallis, C. D., Lee, D., Wexler, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+extra-thoracic+airways+and+deposited+particles+in+laboratory+animals.">Imaging extra-thoracic airways and deposited particles in laboratory animals.</a> J Aerosol Sci. 45, 40-49 (2012).</li><li>Thomas, S. M., Bednarek, J., Janssen, W. J., Hume, P. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Air-inflation+of+murine+lungs+with+vascular+perfusion-fixation.">Air-inflation of murine lungs with vascular perfusion-fixation.</a> J Vis Exp. 168, e62215 (2021).</li><li>Bankhead, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=QuPath:+Open+source+software+for+digital+pathology+image+analysis.">QuPath: Open source software for digital pathology image analysis.</a> Sci Rep. 7 (1), 16878 (2017).</li><li>Davenport, M. L., Sherrill, T. P., Blackwell, T. S., Edmonds, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+and+inflation+of+the+mouse+lung+for+tumor+histology.">Perfusion and inflation of the mouse lung for tumor histology.</a> J Vis Exp. 162, e60605 (2020).</li><li>Hsia, C. C. W., Hyde, D. M., Ochs, M., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+official+research+policy+statement+of+the+American+Thoracic+Society/European+Respiratory+Society:+Standards+for+quantitative+assessment+of+lung+structure.">An official research policy statement of the American Thoracic Society/European Respiratory Society: Standards for quantitative assessment of lung structure.</a> Am J Respir Crit Care Med. 181 (4), 394-418 (2010).</li><li>Gil, J., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular+lining+of+bronchioles+after+perfusion-fixation+of+rat+lungs+for+electron+microscopy.">Extracellular lining of bronchioles after perfusion-fixation of rat lungs for electron microscopy.</a> Anat Rec. 169 (2), 185-199 (1971).</li><li>Bachofen, H., Ammann, A., Wangensteen, D., Weibel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+fixation+of+lungs+for+structure-function+analysis:+credits+and+limitations.">Perfusion fixation of lungs for structure-function analysis: credits and limitations.</a> J Appl Physiol Respir Environ Exerc Physiol. 53 (2), 528-533 (1982).</li><li>Evans, C. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+polymeric+mucin+Muc5ac+is+required+for+allergic+airway+hyperreactivity.">The polymeric mucin Muc5ac is required for allergic airway hyperreactivity.</a> Nat Commun. 6, 6281 (2015).</li><li>Galati, D. F., Asai, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunofluorescence+microscopy.">Immunofluorescence microscopy.</a> Curr Protoc. 3 (8), e842 (2023).</li><li>Hickey, S. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+microscopy-An+outline+of+hardware,+biological+handling,+and+fluorophore+considerations.">Fluorescence microscopy-An outline of hardware, biological handling, and fluorophore considerations.</a> Cells. 11 (1), 35 (2021).</li><li>Risling, T. E., Caulkett, N. A., Florence, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Open-drop+anesthesia+for+small+laboratory+animals.">Open-drop anesthesia for small laboratory animals.</a> Can Vet J. 53 (3), 299-302 (2012).</li><li>Bodnar, M. J., Ratuski, A. S., Weary, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+isoflurane+anesthesia+using+the+drop+method.">Mouse isoflurane anesthesia using the drop method.</a> Lab Anim. 57 (6), 623-630 (2023).</li></ol>

The authors have nothing to disclose.

We have established a pipeline for spore inhalation and analysis of the spatial deposition of the inhaled spores. This pipeline provides valuable information to determine the relevant stromal regions of the lung affected by inhalation of Coccidioides posadasii cts2/ard1/cts3&#916; arthroconidia. We have observed that Coccidioides spores and similarly sized inert particles (data not shown) accumulate in distal bronchiolar regions and broncho-alveolar junctions rather than fully dispersed throughout alve...

Humans can inhale up to billions of spores per day from a variety of environmental fungi1. To understand our barrier defenses against these inhaled spores, we must appreciate the precise microanatomical environments where these spores land within the airways and lung parenchyma. The cellular composition of the airways (i.e., epithelial cells) significantly transforms along the trachea, bronchi, bronchioles, and alveoli. Each of these distinct regions is composed of different cell types with discrete functions which avails an arsenal of defenses to prevent pathologic infection.
The precise location of pulmonary fungal spore deposition can vary between the airway lumens of columnar epithelial-lined bronchioles, alveolar ducts, or alveoli2. Most clinically relevant fungal species produce spores between 1 &#181;m and 10 &#181;m in diameter3. The deposition of these spore particles in the lung depends on several factors, such as aerodynamics, density, electrical charge, and phoretic forces, which can influence the mechanism of sedimentation after inhalation4. Generally, large particles (&#62; 6 &#181;m) deposit in the upper airway, medium-sized particles (2-6 &#181;m) can deposit in smaller airways, and small particles (&#60;2 &#181;m) reach the alveolar region5. Aspergillus spores (2-3 &#181;m) have been reported to reach alveolar spaces, but clinical pathology reports also indicate a significant burden of bronchial and bronchiolar disease6. There is also increasing recognition of endobronchial fungal infections of Aspergillus fumigatus, Coccidioides immitis, Candida species, Cryptococcus neoformans, Histoplasma capsulatum, and Zygomycetes due to the increasing popularity of flexible bronchoscopy7. Recent advances in microscopic imaging of Aspergillus infections in mice have also revealed that more proximal airspaces such as bronchi and bronchioles may bear the highest burden for pathologic fungal proliferation8. Research on the host response to pulmonary fungal infections has shown that both bronchiolar epithelial cells and alveolar epithelial cells play important roles as immune sentinels, so elucidating the exact sites of spore deposition and epithelial interaction will be vital for future work2,9,10.
Studying these proximal airway pathogenesis dynamics is difficult because standard lung fixation and sectioning preparation techniques can displace spores from these proximal positions in epithelial-lined airways and push them toward distal terminal alveolar regions. Commonly, 10% formalin or 4% paraformaldehyde is used to inflate the lungs and rapidly expose the entire lung to fixative. When preparing lungs for cryosectioning, some groups administer optimal cutting temperature (OCT) compound into the lungs to improve cryosectioning performance11. These practices are useful in the right context but have been found by our lab and other groups to displace spores and particles from proximal locations, thereby interfering with interpretations about the spore-exposed cell types and the subsequent host response12.
To accurately establish the microanatomical localization of inhaled fungal spores, we have developed a quick, low-resource method for the preservation of the location of fungal spores in the airways of mice. We adapted a murine air-inflation vascular perfusion-fixation method from Thomas et al. (2021), where we reduced the amount of equipment, time, and technical skill required to achieve a satisfactory result13. From the method described here, we have observed that Coccidioides posadasii cts2/ard1/cts3&#916; arthroconidia (3-5 &#181;m in size) accumulate more proximally than previously shown, namely in distal bronchioles and broncho-alveolar junctions rather than terminal alveoli. This information can focus&#160;biological questions concerning the critical cell types associated with these regions of the lung and their influence on the early responses to inhaled fungal spores.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>18 G, 1 1/2 needle (305185)</td>
			<td>Fisher</td>
			<td>305185</td>
			<td></td>
		</tr>
		<tr>
			<td>1 L&#160; Screwtop Jar&#160; (Nalgene)</td>
			<td>Fisher Scientific</td>
			<td>11-823-33</td>
			<td></td>
		</tr>
		<tr>
			<td>Air-Tite Bulk Unsterile Syringes 10 mL Luer Lock</td>
			<td>Fisher</td>
			<td>14-817-175</td>
			<td></td>
		</tr>
		<tr>
			<td>AnaSed Injection (xylazine sterile solution)</td>
			<td>Akorn</td>
			<td>59399-110-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Animal-Free Blocker and Diluent, R.T.U.</td>
			<td>Vector</td>
			<td>SP-5035-100</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Insyte Autoguard Winged Shielded IV Catheter with BD Vialon Catheter Material 18 G x 1.88 in</td>
			<td>BD</td>
			<td>381547</td>
			<td></td>
		</tr>
		<tr>
			<td>BD Pharmingen Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block&#8482;)</td>
			<td>BD</td>
			<td>553142</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTracker Orange CMRA Dye</td>
			<td>Fisher Scientific</td>
			<td>NC0873640</td>
			<td></td>
		</tr>
		<tr>
			<td>CFSE</td>
			<td>Labviva</td>
			<td>75003</td>
			<td></td>
		</tr>
		<tr>
			<td>Coccidioides posadasii cts2/ard1/cts3&#916;&#160;</td>
			<td>BEI Resources</td>
			<td>NR-166</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning 70 micron strainers</td>
			<td>VWR</td>
			<td>10054-456</td>
			<td></td>
		</tr>
		<tr>
			<td>EpCAM AlexaFluor647 monoclonal antibody</td>
			<td>Biolegend</td>
			<td>118211</td>
			<td></td>
		</tr>
		<tr>
			<td>Exel International HYpodermic Needles 30 G x 1/2&#34;</td>
			<td>Labviva</td>
			<td>EN3012</td>
			<td></td>
		</tr>
		<tr>
			<td>Fisherbrand Sterile Syringes for Single Use (1mL, Leur Slip)</td>
			<td>Fisher</td>
			<td>14-955-462</td>
			<td></td>
		</tr>
		<tr>
			<td>Glucose Monohydrate</td>
			<td>Azer Scientific&#160;</td>
			<td>ES17530-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>High Vacuum Grease</td>
			<td>VWR</td>
			<td>59344-055</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoechst 33342 Solution 20 mM (5 mL)</td>
			<td>ThermoFischer</td>
			<td>62249</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane USP</td>
			<td>Covetrus</td>
			<td>29405</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine Hydrochloride</td>
			<td>Dechra</td>
			<td>1000001250</td>
			<td></td>
		</tr>
		<tr>
			<td>KIMWIPES Delicate Task Wipers (4.4&#39;&#39; x 8.4&#34;)</td>
			<td>VWR</td>
			<td>21905-026</td>
			<td></td>
		</tr>
		<tr>
			<td>Lexer Baby Scissors</td>
			<td>FST</td>
			<td>14078-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-Adson Forceps</td>
			<td>FST</td>
			<td>11018-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Neutral Buffered Formalin (10%) (Azer Scientific)</td>
			<td>Fisher</td>
			<td>22-026-350</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc EasYFlask tissue culture flasks, T75, filter caps</td>
			<td>VWR</td>
			<td>15708-134</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>VWR</td>
			<td>45000-446</td>
			<td></td>
		</tr>
		<tr>
			<td>Peel-A-Way embedding molds</td>
			<td>Sigma</td>
			<td>E6032-1CS</td>
			<td></td>
		</tr>
		<tr>
			<td>QuPath 0.5.1 Software</td>
			<td>Open-source</td>
			<td>https://qupath.github.io/</td>
			<td></td>
		</tr>
		<tr>
			<td>Silk Suture thread size 3/0</td>
			<td>FST</td>
			<td>18020-30</td>
			<td></td>
		</tr>
		<tr>
			<td>SlidesMicro Slides Superfrost Plus&#160;</td>
			<td>VWR</td>
			<td>48311-703</td>
			<td></td>
		</tr>
		<tr>
			<td>SlowFade Glass Soft-set Antifade Mountant (2 mL)</td>
			<td>ThermoFischer</td>
			<td>S36917</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma</td>
			<td>S0389-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue-Tek O.C.T. Compound, Sakura Finetek</td>
			<td>VWR</td>
			<td>25608-930</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween 20</td>
			<td>ThermoFischer</td>
			<td>J20605.AP</td>
			<td></td>
		</tr>
		<tr>
			<td>Vector Laboratories ImmEDGE Hydrophobic Barrier Pen Set Of 2</td>
			<td>Fisher Scientific</td>
			<td>NC9545623</td>
			<td></td>
		</tr>
		<tr>
			<td>VWR Micro Cover Glasses, Rectangular (24 mm x 40 mm #1.5)</td>
			<td>VWR</td>
			<td>48393-230</td>
			<td></td>
		</tr>
		<tr>
			<td>White Plastic Wire Mesh</td>
			<td>MAPORCH</td>
			<td>789862904922</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast Extract</td>
			<td>Fisher</td>
			<td>BP9727-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss AxioScan 7&#160;</td>
			<td>Carl Zeiss Microscopy GmbH</td>
			<td>https://www.zeiss.com/microscopy/us/products/imaging-systems/axioscan-for-biology.html</td>
			<td>multichannel fluorescent microscope</td>
		</tr>
		<tr>
			<td>ZEN 3.7 Software</td>
			<td>Carl Zeiss Microscopy GmbH</td>
			<td>https://www.zeiss.com/microscopy/us/products/software/zeiss-zen.html</td>
			<td>microscopy software</td>
		</tr>
	</tbody>
</table>

efficient method for imaging murine lungs that preserves spatial dynamics of fungal spores in the airways

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and bronchioles, branch until terminating in the alveolar airspace where gas exchange occurs. Infections originating in the bronchioles or alveoli elicit distinct host responses and disease manifestations. Therefore, understanding precisely where spores naturally localize in the lungs, particularly soon after infection, expands opportunities for investigation of host-pathogen interactions. Herein, we detail an in-situ analysis of lungs from mice infected with Coccidioides posadasii cts2/ard1/cts3&#916; arthroconidia. Conventional methods for histological preservation involve liquid inflation of the airways with a fixative solution, which displaces the natural location of aspirated fungal particles, pushing spores from proximal bronchioles to terminal airspaces.
Conversely, this method of air-inflation with blood vasculature perfusion-fixation preserves the physiologic position of fungal spores within the bronchioles. Moreover, we describe a simple approach to cryopreserving, embedding, and imaging lung specimens. We also share high-throughput computational techniques via the open-source QuPath program to analyze the spatial distribution of fungal spores within the lung. The method presented here is simple and quick, requires minimal equipment to perform, and can be easily adapted for use with many respiratory fungal infection models.

This method ultimately produces immunofluorescent images of mouse lungs using physiologic air inflation to leave the airways undisturbed. Importantly, multiple checkpoints along the way will confirm that components of the protocol have been performed successfully. During the inoculation, it is important to confirm that the inoculum was aspirated by feeling for &#34;crackles&#34; on the posterior chest wall of the mouse that indicate the liquid has entered the airways. If there is no sensation of crackles, it is possible ...

Efficient Method for Imaging Murine Lungs that Preserves Spatial Dynamics of Fungal Spores in the Airways

We present a method for a fungal pathogenesis model that preserves the natural positioning of fungal spores in the lung airways for analysis via fluorescent microscopy.

Fungi infect humans when environmental spores are inhaled into the lungs. The lung is a heterogeneous organ. Conducting airways, including bronchi and ...

efficient-method-for-imaging-murine-lungs-that-preserves-spatial

Research

JoVE Journal

Immunology and Infection

244 Views.  Rutgers - The State University of New Jersey. We present a method for a fungal pathogenesis model that preserves the natural positioning of fungal spores in the lung airways for analysis via fluorescent microscopy. 

Video: Efficient Method for Imaging Murine Lungs that Preserves Spatial Dynamics of Fungal Spores in the Airways

Establishment of an In vitro System to Study Intracellular Behavior of Candida glabrata in Human THP-1 Macrophages

A cell culture model system, if a close mimic of host environmental conditions, can serve as an inexpensive, reproducible and easily manipulatable alternative to animal model systems for the study of a specific step of microbial pathogen infection. A human monocytic cell line THP-1 which, upon phorbol ester treatment, is differentiated into macrophages, has previously been used to study virulence strategies of many intracellular pathogens including Mycobacterium tuberculosis. Here, we discuss a protocol to enact an in vitro cell culture model system using THP-1 macrophages to delineate the interaction of an opportunistic human yeast pathogen Candida glabrata with host phagocytic cells. This model system is simple, fast, amenable to high-throughput mutant screens, and requires no sophisticated equipment. A typical THP-1 macrophage infection experiment takes approximately 24 hr with an additional 24-48 hr to allow recovered intracellular yeast to grow on rich medium for colony forming unit-based viability analysis. Like other in vitro model systems, a possible limitation of this approach is difficulty in extrapolating the results obtained to a highly complex immune cell circuitry existing in the human host. However, despite this, the current protocol is very useful to elucidate the strategies that a fungal pathogen may employ to evade/counteract antimicrobial response and survive, adapt, and proliferate in the nutrient-poor environment of host immune cells.

Establishment of an In vitro System to Study Intracellular Behavior of Candida glabrata in Human THP-1 Macrophages

The current article outlines a protocol to establish an in vitro cell culture model system to study the interaction of a facultative intracellular human fungal pathogen Candida glabrata with human macrophages which will be a useful tool to advance our knowledge of fungal virulence mechanisms.

A cell culture model system, if a close mimic of host environmental conditions, can serve as an inexpensive, reproducible and easily manipulatable ...

A Reference Broth Microdilution Method for Dalbavancin In Vitro Susceptibility Testing of Bacteria that Grow Aerobically

Antimicrobial susceptibility testing (AST) is performed to assess the in vitro activity of antimicrobial agents against various bacteria. The AST results, which are expressed as minimum inhibitory concentrations (MICs) are used in research for antimicrobial development and monitoring of resistance development and in the clinical setting for antimicrobial therapy guidance. Dalbavancin is a semi-synthetic lipoglycopeptide antimicrobial agent that was approved in May 2014 by the Food and Drug Administration (FDA) for the treatment of acute bacterial skin and skin structure infections caused by Gram-positive organisms. The advantage of dalbavancin over current anti-staphylococcal therapies is its long half-life, which allows for once-weekly dosing. Dalbavancin has activity against Staphylococcus aureus (including both methicillin-susceptible S. aureus [MSSA] and methicillin-resistant S. aureus [MRSA]), coagulase-negative staphylococci, Streptococcus pneumoniae, Streptococcus anginosus group, &#946;-hemolytic streptococci and vancomycin susceptible enterococci. Similar to other recent lipoglycopeptide agents, optimization of CLSI and ISO broth susceptibility test methods includes the use of dimethyl sulfoxide (DMSO) as a solvent when preparing stock solutions and polysorbate 80 (P80) to alleviate adherence of the agent to plastic. Prior to the clinical studies and during the initial development of dalbavancin, susceptibility studies were not performed with the use of P-80 and MIC results tended to be 2-4 fold higher and similarly higher MIC results were obtained with the agar dilution susceptibility method. Dalbavancin was first included in CLSI broth microdilution methodology tables in 2005 and amended in 2006 to clarify use of DMSO and P-80. The broth microdilution (BMD) procedure shown here is specific to dalbavancin and is in accordance with the CLSI and ISO methods, with step-by-step detail and focus on the critical steps added for clarity.

A Reference Broth Microdilution Method for Dalbavancin In Vitro Susceptibility Testing of Bacteria that Grow Aerobically

Here, we present a protocol for determination of dalbavancin susceptibility of clinically relevant Gram-positive bacteria using a broth microdilution reference methodology.

Antimicrobial susceptibility testing (AST) is performed to assess the in vitro activity of antimicrobial agents against various bacteria. The AST results, ...

An Endothelial Planar Cell Model for Imaging Immunological Synapse Dynamics

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological synapses&#39;. Within these intimate cell-cell interfaces discrete sub-cellular clusters of MHC/Ag-TCR, F-actin, adhesion and signaling molecules form and remodel rapidly. These dynamics are thought to be critical determinants of both the efficiency and quality of the immune responses that develop and therefore of protective versus pathologic immunity. Current understanding of immunological synapses with physiologic APCs is limited by the inadequacy of the obtainable imaging resolution. Though artificial substrate models (e.g., planar lipid bilayers) offer excellent resolution and have been extremely valuable tools, they are inherently non-physiologic and oversimplified. Vascular and lymphatic endothelial cells have emerged as an important peripheral tissue (or stromal) compartment of &#39;semi-professional APCs&#39;. These APCs (which express most of the molecular machinery of professional APCs) have the unique feature of forming virtually planar cell surface and are readily transfectable (e.g., with fluorescent protein reporters). Herein a basic approach to implement endothelial cells as a novel and physiologic &#39;planar cellular APC model&#39; for improved imaging and interrogation of fundamental antigenic signaling processes will be described.

Adaptive immunity is controlled by dynamic 'immunological synapses' formed between T cells and antigen presenting cells. This protocol describes methods for investigating endothelial cells both as understudied physiologic APCs and as a novel type of 'planar cellular APC model'.

Adaptive immunity is regulated by dynamic interactions between T cells and antigen presenting cells (&#39;APCs&#39;) referred to as &#39;immunological ...

An Efficient and High Yield Method for Isolation of Mouse Dendritic Cell Subsets

Dendritic cells (DCs) are professional antigen-presenting cells primarily responsible for acquiring, processing and presenting antigens on antigen presenting molecules to initiate T-cell-mediated immunity. Dendritic cells can be separated into several phenotypically and functionally heterogeneous subsets. Three important subsets of splenic dendritic cells are plasmacytoid, CD8&#945;Pos and CD8&#945;Neg cells. The plasmacytoid DCs are natural producers of type I interferon and are important for anti-viral T cell immunity. The CD8&#945;Neg DC subset is specialized for MHC class II antigen presentation and is centrally involved in priming CD4 T cells. The CD8&#945;Pos DCs are primarily responsible for cross-presentation of exogenous antigens and CD8 T cell priming. The CD8&#945;Pos DCs have been demonstrated to be most efficient at the presentation of glycolipid antigens by CD1d molecules to a specialized T cell population known as invariant natural killer T (iNKT) cells. Administration of Flt-3 ligand increases the frequency of migration of dendritic cell progenitors from bone marrow, ultimately resulting in expansion of dendritic cells in peripheral lymphoid organs in murine models. We have adapted this model to purify large numbers of functional dendritic cells for use in cell transfer experiments to compare in vivo proficiency of different DC subsets.

Distinct dendritic cell subsets exist as rare populations in lymphoid organs, and therefore are challenging to isolate in sufficient numbers and purity for immunological experiments. Here we describe a high efficiency, high yield method for isolation of all of the currently known major subsets of mouse splenic dendritic cells.

Dendritic cells (DCs) are professional antigen-presenting cells primarily responsible for acquiring, processing and presenting antigens on antigen ...

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo Cell Trafficking by PET/CT

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte cell culture and 64Cu-antibody receptor targeting of the cells. In vitro evaluation of the radiolabel and non-invasive in vivo cell tracking in an animal model of an airway delayed-type hypersensitivity reaction (DTHR) by PET/CT are described.
In detail, the conjugation of a mAb with the chelator 1,4,7,10-tetraazacyclododecane-1,4,7,10-tetraacetic acid (DOTA) is shown. Following the production of radioactive 64Cu, radiolabeling of the DOTA-conjugated mAb is described. Next, the expansion of chicken ovalbumin (cOVA)-specific CD4+ interferon (IFN)-&#947;-producing T helper cells (cOVA-TH1) and the subsequent radiolabeling of the cOVA-TH1 cells are depicted. Various in vitro techniques are presented to evaluate the effects of 64Cu-radiolabeling on the cells, such as the determination of cell viability by trypan blue exclusion, the staining for apoptosis with Annexin V for flow cytometry, and the assessment of functionality by IFN-&#947; enzyme-linked immunosorbent assay (ELISA). Furthermore, the determination of the radioactive uptake into the cells and the labeling stability are described in detail. This protocol further describes how to perform cell tracking studies in an animal model for an airway DTHR and, therefore, the induction of cOVA-induced acute airway DHTR in BALB/c mice is included. Finally, a robust PET/CT workflow including image acquisition, reconstruction, and analysis is presented.
The 64Cu-antibody receptor targeting approach with subsequent receptor internalization provides high specificity and stability, reduced cellular toxicity, and low efflux rates compared to common PET-tracers for cell labeling, e.g.64Cu-pyruvaldehyde bis(N4-methylthiosemicarbazone) (64Cu-PTSM). Finally, our approach enables non-invasive in vivo cell tracking by PET/CT with an optimal signal-to-background ratio for 48 h. This experimental approach can be transferred to different animal models and cell types with membrane-bound receptors that are internalized.

Murine Lymphocyte Labeling by 64Cu-Antibody Receptor Targeting for In Vivo  Cell Trafficking by PET/CT

Following the preparation of a 64Cu-modified monoclonal antibody binding to a transgenic murine T cell receptor, T cells are radiolabeled in vivo, analyzed for viability, functionality, labeling stability and apoptosis, and adoptively transferred into mice with an airway delayed-type hypersensitivity reaction for non-invasive imaging by positron emission tomography/computed tomography (PET/CT).

This protocol illustrates the production of 64Cu and the chelator conjugation/radiolabeling of a monoclonal antibody (mAb) followed by murine lymphocyte ...

Bronchoalveolar Lavage of Murine Lungs to Analyze Inflammatory Cell Infiltration

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain insight into an ongoing disease state.
Here, a simple and efficient method is described to perform BAL on murine lungs without the need of special tools or equipment. BAL fluid is isolated by inserting a catheter in the trachea of terminally anesthetized mice, through which a saline solution is instilled into the bronchioles. The instilled fluid is gently retracted to maximize BAL fluid retrieval and to minimize shearing forces. This technique allows the viability, function, and structure of cells within the airways and BAL fluid to be preserved.
Numerous techniques may be applied to gain further understanding of the disease state of the lung. Here, a commonly used technique for the identification and enumeration of different types of immune cells is described, where&#160;flow cytometry is combined with a select panel of fluorescently labeled cell surface-specific markers. The BAL procedure presented here can also be used to analyze infectious agents, fluid constituents, or inhaled particles within murine lungs.

The health status of the lung is reflected by the type and number of immune cells that are present in the bronchioles of the lung. We describe a bronchoalveolar lavage technique that allows the isolation and study of nonadherent cells and soluble factors from the lower respiratory tract of mice.

Bronchoalveolar Lavage (BAL) is an experimental procedure that is used to examine the cellular and acellular content of the lung lumen ex vivo to gain ...

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research investigating immunological responses against A. fumigatus has been limited by the lack of consistent and reliable assays for measuring the antifungal activity of specific immune cells in vitro. A new method is described to assess the antifungal activity of primary monocytes and neutrophils from human donors against A. fumigatus using FLuorescent Aspergillus REporter (FLARE) conidia. These conidia contain a genetically encoded dsRed reporter, which is constitutively expressed by live FLARE conidia, and are externally labeled with Alexa Fluor 633, which is resistant to degradation within the phagolysosome, thus allowing a distinction between live and dead A. fumigatus conidia. Video microscopy and flow cytometry are subsequently used to visualize the interaction between conidia and innate immune cells, assessing fungicidal activity whilst also providing a wealth of information on phagocyte migration, phagocytosis and the inhibition of fungal growth. This novel technique has already provided exciting new insights into the host-pathogen interaction of primary immune cells against A. fumigatus. It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.
It is important to note the laboratory setup required to perform this assay, including the necessary microscopy and flow cytometry facilities, and the capacity to work with human donor blood and genetically manipulated fungi. However, this assay is capable of generating large amounts of data and can reveal detailed insights into the antifungal response. This protocol has successfully been used to study the host-pathogen interaction of primary immune cells against A. fumigatus.

Live Imaging of Antifungal Activity by Human Primary Neutrophils and Monocytes in Response to A. fumigatus

Here, we describe a protocol to assess antifungal activity of primary human immune cells in real-time using fluorescent Aspergillus reporter conidia in conjunction with live-cell video microscopy and flow cytometry. Generated data provide insight into host cell-Aspergillus interactions such as fungicidal activity, phagocytosis, cell migration and inhibition of fungal growth.

Aspergillus fumigatus is an opportunistic fungal pathogen causing invasive infections in immunocompromised hosts with a high case-fatality rate. Research ...

Intravital Imaging of Intraepithelial Lymphocytes in Murine Small Intestine

Intraepithelial lymphocytes expressing &#947;&#948; T cell receptor (&#947;&#948; IEL) play a key role in immune surveillance of the intestinal epithelium. Due in part to the lack of a definitive ligand for the &#947;&#948; T cell receptor, our understanding of the regulation of &#947;&#948; IEL activation and their function in vivo remains limited. This necessitates the development of alternative strategies to interrogate signaling pathways involved in regulating &#947;&#948; IEL function and the responsiveness of these cells to the local microenvironment. Although &#947;&#948; IELs are widely understood to limit pathogen translocation, the use of intravital imaging has been critical to understanding the spatiotemporal dynamics of IEL/epithelial interactions at steady-state and in response to invasive pathogens. Herein, we present a protocol for visualizing IEL migratory behavior in the small intestinal mucosa of a GFP &#947;&#948; T cell reporter mouse using inverted spinning disk confocal laser microscopy. Although the maximum imaging depth of this approach is limited relative to the use of two-photon laser-scanning microscopy, spinning disk confocal laser microscopy provides the advantage of high speed image acquisition with reduced photobleaching and photodamage. Using 4D image analysis software, T cell surveillance behavior and their interactions with neighboring cells can be analyzed following experimental manipulation to provide additional insight into IEL activation and function within the intestinal mucosa.

We describe a method to visualize GFP-labeled &#947;&#948; IELs using intravital imaging of murine small intestine by inverted spinning disk confocal microscopy. This technique enables the tracking of live cells within the mucosa for up to 4 h and can be used to investigate a variety of intestinal immune-epithelial interactions. &#160;

Intraepithelial lymphocytes expressing &#947;&#948; T cell receptor (&#947;&#948; IEL) play a key role in immune surveillance of the intestinal ...

Highly Efficient Transfection of Primary Macrophages with In Vitro Transcribed mRNA

Macrophages are phagocytic cells specialized in detecting molecules of non-self origin. To this end, they are equipped with a large array of pattern recognition receptors (PRRs). Unfortunately, this also makes macrophages particularly challenging to transfect as the transfection reagent and the transfected nucleic acids are often recognized by the PRRs as non-self. Therefore, transfection often results in macrophage activation and degradation of the transfected nucleic acids or even in suicide of the macrophages. Here, we describe a protocol that allows highly efficient transfection of murine primary macrophages such as peritoneal macrophages (PM) and bone marrow-derived macrophages (BMDM) with mRNA in vitro transcribed from DNA templates such as plasmids. With this simple protocol, transfection rates of about 50-65% for PM and about 85% for BMDM are achieved without cytotoxicity or immunogenicity observed. We describe in detail the generation of mRNA for transfection from DNA constructs such as plasmids and the transfection procedure.

Macrophages, especially primary macrophages, are challenging to transfect as they specialize in detecting molecules of non-self origin. We describe a protocol that allows highly efficient transfection of primary macrophages with mRNA generated from DNA templates such as plasmids.

Macrophages are phagocytic cells specialized in detecting molecules of non-self origin. To this end, they are equipped with a large array of pattern ...

Optimization of the Cuff Technique for Murine Heart Transplantation

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to improve surgical efficiency. In our lab, we have optimized the cuff technique with two major steps. First, we used the inner tube technique to insert a temporary inner tube into the external jugular vein and carotid artery blood vessel to facilitate eversion of the vessel over the cuff. Second, we performed complete heterotopic cardiac transplantation through the collaboration of two experienced surgeons. These modifications effectively reduced the operation time to 25 minutes, with a success rate of 95%. In this report, we describe these procedures in detail and provide a supplemental video. We believe that this report on the improved cuff technique will offer practical guidance for murine heterotopic heart transplantation and will enhance the utility of this mouse model for basic research.

We introduce an inner tube approach to the cuff technique for mouse cervical heterotopic heart transplantation to help evert the vessel over the cuff. We found that cooperation between two experienced surgeons markedly shortens the operation time.

Murine cardiac transplantation has been performed for more than 40 years. With advancements in microsurgery, certain new techniques have been used to ...